Anim. Reprod., v.12, n.1, p.93-104, Jan./Mar. 2015
Germline modification of domestic animals
L. Tang, R. González, I.Dobrinski1
Department of Comparative Biology and Experimental Medicine, Faculty of Veterinary Medicine,
University of Calgary, Calgary, AB, Canada.
Abstract
Genetically-modified domestic animal models
are of increasing significance in biomedical research
and agriculture. As authentic ES cells derived from
domestic animals are not yet available, the prevailing
approaches for engineering genetic modifications in
those animals are pronuclear microinjection and somatic
cell nuclear transfer (SCNT, also known as cloning).
Both pronuclear microinjection and SCNT are
inefficient, costly, and time-consuming. In animals
produced by pronuclear microinjection, the exogenous
transgene is usually inserted randomly into the genome,
which results in highly variable expression patterns and
levels in different founders. Therefore, significant
efforts are required to generate and screen multiple
founders to obtain animals with optimal transgene
expression. For SCNT, specific genetic modifications
(both gain-of-function and loss-of-function) can be
engineered and carefully selected in the somatic cell
nucleus before nuclear transfer. SCNT has been used to
generate a variety of genetically modified animals such
as goats, pigs, sheep and cattle; however, animals
resulting from SCNT frequently suffer from
developmental abnormalities associated with incomplete
nuclear reprogramming. Other strategies to generate
genetically-modified animals rely on the use of the
spermatozoon as a natural vector to introduce genetic
material into the female gamete. This sperm mediated
DNA
transfer
(SMGT)
combined
with
intracytoplasmatic sperm injection (ICSI) has relatively
high efficiency and allows the insertion of large DNA
fragments, which, in turn, enhance proper gene
expression. An approach currently being developed to
complement SCNT for producing genetically modified
animals is germ cell transplantation using genetically
modified male germline stem cells (GSCs). This
approach relies on the ability of GSCs that are
genetically modified in vitro to colonize the recipient
testis and produce donor derived sperm upon
transplantation. As the genetic change is introduced into
the male germ line just before the onset of
spermatogenesis, the time required for the production of
genetically modified sperm is significantly shorter using
germ cell transplantation compared to cloning or
embryonic stem (ES) cell based technology. Moreover,
the GSC-mediated germline modification circumvents
_________________________________________
1
Corresponding author: [email protected]
Phone: +1(403)210-6523; Fax: +1(403)210-7882
Received: May 30, 2014
Accepted: August 8, 2014
problems associated with embryo manipulation and
nuclear reprogramming. Currently, engineering targeted
mutations in domestic animals using GSCs remains a
challenge as GSCs from those animals are difficult to
maintain in vitro for an extended period of time. Recent
advances in genome editing techniques such as ZincFinger Nucleases (ZFNs), Transcription Activator-like
Effector Nucleases (TALENs) and Clustered Regularly
Interspaced Short Palindromic Repeats (CRISPRs)
greatly enhance the efficiency of engineering targeted
genetic change in domestic animals as demonstrated by
the generation of several gene knock-out pig and cattle
models using those techniques. The potential of GSCmediated germline modification in making targeted
genetic modifications in domestic animal models will be
maximized if those genome editing techniques can be
applied in GSCs.
Keywords: genetic engineering, large animal models,
male germline, transgenesis.
Introduction
Genetic manipulation and public acceptance
Genetic modification of an animal involves
altering its genetic material by adding, changing or
removing certain DNA sequences in a way that does not
occur naturally (European Food Safety Authority;
www.efsa.europa.eu). While the selection of certain
animal traits has been achieved by selective breeding
since the beginning of animal domestication, the advent
of molecular techniques has allowed to shortcut the long
process of natural breeding for editing animal genomes.
Genetically-modified domestic animal models are of
increasing significance in biomedical research and
agriculture. More progress has been made in the field of
biomedicine than for agricultural purposes due to better
public acceptance of biotechnology when the intention
is to generate health benefits, followed by benefits for
the environment but more concern is generated by food
technology (Einsiedel, 2005). The public opinion also
reflected more objections to the genetic manipulation of
animals than plants or microorganisms (Einsiedel, 2005).
It is expected that the number of genetically modified
animals and their applications will increase in the near
future and so it will the public concern and awareness,
which influences policies by the regulatory agencies.
Tang et al. Germline modification of domestic animals.
The regulatory agencies for genetically modified
animals include the United States Food and Drug
Administration (FDA; www.fda.gov), the European
Medicine Agency (EMA; www.ema.europa.eu); the
European
Food
Safety
Authority
(EFSA;
www.efsa.europa.eu) and the Food and Agriculture
Organization of the United Nations (FAO)/World
Health Organization (WHO).
Use of large animal models in Biomedicine and
Agriculture
The availability of suitable animal models is
essential for the development of new therapies.
Certainly the use of rodents has been very useful in
understanding the function of many genes by means of
targeted mutagenesis. However, although they can be
very useful for the development of new technologies,
rodent models do not always faithfully reflect the
patient situation and many scientific discoveries in
rodent models do not translate into the clinical situation.
Large animals are genetically outbred and more
closely related to humans than rodents and the pig has
become a very good model for translational biomedical
research because it resembles humans more closely than
rodents from a physiologic, anatomic and genetic point
of view (Aigner et al., 2010; Bode et al., 2010; Luo et
al., 2012). Genetically modified domestic animals are
currently available as models for diabetes, cystic
fibrosis, cardiovascular diseases and neurodegenerative
diseases, among others (Aigner et al., 2010; Luo et al.,
2012). Other applications of farm animals include their
use for xenotransplantation, as bioreactors for the
production of enzymes, hormones, antibodies or other
compounds of pharmaceutical interest (Niemann and
Kues, 2007; Tan et al., 2012). In the field of agriculture,
genetically modified animals can result in improved
production traits, resistance to diseases, improved
adaptation to the production systems or the environment
and the creation of more environmentally friendly
livestock (Golovan et al., 2001; Donovan et al., 2005;
Niemann and Kues, 2007; Richt et al., 2007; Aigner et
al., 2010; Kues and Niemann, 2011; Piedrahita and
Olby, 2011; Luo et al., 2012). However, although the
first genetically modified animals were generated over
three decades ago (Gordon and Ruddle, 1981; Hammer
et al., 1985), the technology has not yet fulfilled the
original expectations. So far, the European Medicine
Agency approved in 2006 the first biological product
derived from genetically engineered animals, ATryn, an
alternative to antithrombin derived from human plasma
(GTC Biotherapeutics UK Limited). The same product
was later authorized for marketing by the US FDA in
2009.
The knowledge at the molecular level has
greatly increased in recent years for domestic species
which certainly has helped making advancements in the
field. Moreover, the biotechnology for generating
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genetically modified animals is currently available, but
especially in the case of livestock, for many laboratories,
the cost is still extremely prohibitive (Kind and
Schnieke, 2008). Even though, the costs of establishing
a cell-based manufacture are greater than the costs for
the development of genetically modified animals (Kind
and Schnieke, 2008). Considering that many of the cell
culture-derived products are retired during phase 1 or
during preclinical investigations (Kind and Schnieke,
2008), it would be useful to invest in generating
genetically engineered animals.
Traditional approaches to generate genetically
modified animals
Pronuclear (PN) injection
Pioneer experiments demonstrated that foreign
DNA can be introduced into fertilized oocytes by
microinjection of DNA molecules into one of the
pronuclei (Gordon et al., 1980; Gordon and Ruddle,
1981; Hammer et al., 1985). Using this technique,
genetically modified rabbits, sheep and pigs were
generated harbouring the MT-hGH gene (Hammer et al.,
1985). However, PN microinjection is a technically
challenging option that requires highly trained personnel.
Moreover, the opacity of the ooplasm in domestic
species is a key feature that contributes technical
difficulties to the detection of the PN and thus, to the
efficiency of the procedure. One of the main limitations
of PN injection is the low efficiency, reaching success
rates of 1-2% or up to 5-10% in livestock in comparison
to up to 20-25% in mice (Clark, 2002; Garrels et al.,
2011). Another drawback is the variable expression of
the transgene due to random integration into the genome
and the variability in the number of inserted copies. The
random integration can cause position effect variegation
which can lead to gene silencing (Opsahl et al., 2002;
Ohtsuka et al., 2012). Despite of the inherent
disadvantages of PN, this approach prevails as one of
the most commonly used methods for generating
genetically modified mice. Although PN injection has
been successfully used to generate genetically modified
livestock (Robl et al., 2007; Luo et al., 2012), the high
cost of screening and maintaining founders in addition
to the inefficiency of the technique, have prevented its
use on a large scale in livestock or other domestic
species. Recent approaches employing PN injection
with the implementation of recombinases or integrases
for targeted mutagenesis have been described holding
more promise than the conventional PN injection-based
transgenesis (Ohtsuka et al., 2012).
Embryonic stem (ES) cells-based targeted transgenesis
Embryonic stem cells are derived from the
inner cell mass of the blastocyst. ES cells are pluripotent
and can be maintained in vitro in an undifferentiated
Anim. Reprod., v.12, n.1, p.93-104, Jan./Mar. 2015
Tang et al. Germline modification of domestic animals.
stage.ES cells can be genetically modified in vitro and
injected to an embryo where ES cells contribute to all
cell types including the germ line.
ES cell-based targeted mutagenesis can
overcome the aforementioned position effect
variegation and repeat-induced gene silencing. With the
use of ES cells, a single copy of the transgene is
integrated into a determined locus by homologous
recombination (HR) allowing the generation of knockouts, knock-ins or the exchange of genes or large
chromosomal regions (Capecchi, 2005; Laible and
Alonso-Gonzalez, 2009; Ohtsuka et al., 2012).
Unfortunately, this technology is only available in mice.
Despite many efforts by multiple laboratories to
generate ES cells in livestock, to date, there are no true
established ES cell lines. So far, the ES-like cells
generated in domestic species do not fulfill the more
stringent in vivo test of chimera formation with germline
transmission (Nowak-Imialek et al., 2011; Piedrahita
and Olby, 2011).
An important breakthrough was the
reprogramming of terminally differentiated fibroblasts
into the pluripotent state by ectopic expression of
several transcription factors, leading to the
establishment of induced pluripotent stem (iPS) cells
(Takahashi and Yamanaka, 2006; Takahashi et al.,
2007). The iPS cells share common characteristics with
ES cells, but they differ in their epigenetic signature and
they display epigenetic memory (Nowak-Imialek et al.,
2011). Given their great potential in regenerative
medicine, gene targeting has been successfully
performed in iPS cells to correct disease related genetic
mutations (Zou et al., 2009; Yusa et al., 2011). The
generation of iPS cells has been reported in several nonrodent species, but their germ line competence has not
been described so far (Nowak-Imialek et al., 2011).
Cloning: somatic cell nuclear transfer (SCNT) and
handmade cloning (HMD)
The advent of new technologies to clone
animals by the transfer of nuclei from somatic cells
(Campbell et al., 1996; Wilmut et al., 1997) opened
new avenues to generate genetically modified livestock
animals. Somatic cell nuclear transfer involves the
enucleation of matured oocytes, followed by the
injection or fusion of the donor cell and activation of the
reconstructed embryo. It has been successful in many
species, including laboratory, domestic and wildlife
species (Galli et al., 2012). The technical and biological
aspects for successful SCNT have been reviewed
elsewhere (Galli et al., 2012). Once SCNT became
available, the first genetically modified animals were
generated shortly afterwards (Schnieke et al., 1997) and
the first knock out livestock was generated by HR in
fibroblasts (McCreath et al., 2000; Denning et al., 2001).
Currently, SCNT is the preferred approach for
generating genetically modified large animals (Schnieke
Anim. Reprod., v.12, n.1, p.93-104, Jan./Mar. 2015
et al., 1997; McCreath et al., 2000; Wang et al., 2008;
Yin et al., 2008; Gomez et al., 2009; Hong et al., 2009;
Kim et al., 2011; Yang et al., 2011a; Jeong et al., 2012;
Luo et al., 2012; Zhang et al., 2014). Without the
establishment of true ES cells in livestock, gene
targeting has been done primarily in fibroblasts for
subsequent SCNT. Somatic cells, however, have a low
frequency of HR in comparison to mouse ES cells and
have a limited life span in culture which hampers the
establishment of cell lines with the desired genetic
modification. Furthermore, transfection of primary cells
might affect their growth and induce senescence before
they can be fully characterized (Laible and AlonsoGonzalez, 2009). Some strategies have been used in an
attempt to circumvent these problems, such as the
introduction of the human telomerase catalytic subunit
(hTERT) to increase the longevity of the primary
somatic cells, but it appears that the constant expression
of hTERT is not compatible with the production of live
animals by SCNT (Laible and Alonso-Gonzalez, 2009).
Other options aim to rejuvenate genetically modified
cell lines by serial cloning, re-deriving the cell lines
from cloned fetuses (Robl et al., 2007; Laible and
Alonso-Gonzalez, 2009), but the rejuvenation process
can increase the accumulation of epigenetic/genetic
errors that eventually may lead to loss of the cloning
potential (Laible and Alonso-Gonzalez, 2009; Liu et al.,
2011). Sequential targeting has been done successfully
in cattle (Kuroiwa et al., 2004) and calves have been
obtained after four rounds of genetic modifications
(Robl et al., 2007).
Somatic cell nuclear transfer requires
micromanipulation, and therefore, highly skilled
technicians and sophisticated equipment. One of the
main drawbacks of the technique is the low
developmental competence of the reconstructed
embryos to produce normal offspring. Usually, SCNTgenerated embryos are associated with higher pregnancy
losses than their in vitro fertilized counterparts, due to
placental abnormalities. However, the cloned animals
that survive to adulthood are normal and are able to
reproduce. The abnormalities observed in SCNTproduced animals are mainly of epigenetic origin and
have been reviewed by others (Galli et al., 2012).
A simplified methodology for cloning by
SCNT, namely, hand-made cloning (HMC), has been
proposed (Vajta et al., 2005) in order to avoid
micromanipulation and its inherent difficulties to make it a
more user-friendly system. With this technique, the zona
pellucida is first digested and the nucleus of the oocyte is
removed using blades. The enucleated cytoplast is
afterwards electrofused with the desired somatic cell to
make the reconstructed embryo. HMC allows the fusion of
several cytoplasts to increase the volume of the
reconstructed embryo which increases the average
number of cells in the resulting blastocysts and thus,
their developmental competence. Important progress has
been made improving blastocyst yield and pregnancy
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Tang et al. Germline modification of domestic animals.
rates after HMC, genetically modified animals have
been generated by this method, such as pigs (Kragh et
al., 2009; Luo et al., 2011, 2012) and sheep (Zhang et
al., 2013) and it is expected that this approach will
become more used in the near future.
Transgenesis through the male germ cell line
Spermatogenesis is a very complex and
coordinated process that occurs in the testis, by which
spermatogonial stem cells undergo mitotic and meiotic
divisions and differentiate leading to the continuous
production of spermatozoa throughout the reproductive
life of a male (Kerr et al., 2006). Spermatogenesis is an
extremely efficient process, therefore, manipulations of
the male germ line provides an attractive approach for
the generation of genetically modified animals.
Different spermatogenic cell types have been targeted
for introducing genetic manipulations in the male germ
line. Mainly the end product of spermatogenesis, the
male gamete that eventually will be used for in vitro or
in vivo fertilization; or the stem cells that will be able to
establish
donor-derived
spermatogenesis
after
transplantation into the testes of recipient animals.
Sperm mediated gene transfer (SMGT) and
intracytoplasmic sperm injection mediated gene transfer
(ICSI-MGT)
The option of SMGT relies on the use of the
spermatozoon as a natural vector for transferring genetic
material into the oocyte. The mechanism of action is
controversial, but exogenous DNA can specifically bind
to the head of the spermatozoon through binding
proteins (Parrington et al., 2011) which are normally
blocked by inhibitory factors (Carballada and Esponda,
2001) in the seminal plasma. Different options have
been used in combination with SMGT to increase its
efficiency, such as liposomes, electroporation or
restriction enzyme mediated integration (REMI;
Parrington et al., 2011). Advantages of SMGT are its
low cost and simplicity, together with the possibility of
harvesting high numbers of spermatozoa, thus, being
feasible in numerous species. The main disadvantages
of this passive technique are the relatively low
efficiency, random integration of DNA and mosaicism.
This approach provoked interest in the scientific
community for its simplicity, but has not been widely
adopted for creating genetically modified animals for its
variable outcomes. The use of SMGT alone or in
combination with other methods to facilitate the
introduction of exogenous DNA through the membrane
of the sperm resulted in the production of transgenic
animals (Wang et al., 2001; Zhang et al., 2012)
Another approach to overcome the difficulties
of PN microinjection in large animals is the use of
intracytoplasmic sperm injection mediated gene transfer
(ICSI-GMT), which can also be considered as an
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extension of SMGT. The procedure is usually carried
out using spermatozoa treated with a detergent, NaOH,
lysolecithin, after freeze-thawing or other methods that
cause damage to the membrane of the spermatozoa
(Moisyadi et al., 2009; Parrington et al., 2011).
Afterwards, the spermatozoa are incubated with the
DNA and the spermatozoon-DNA complex is injected
into matured (metaphase II) oocytes. This tool has
mainly been used in mice (Moisyadi et al., 2009), but it
has been also successfully applied to the pig (Umeyama
et al., 2009). The main drawback of ICSI-MGT is that is
technically complex, requiring skillful and trained
personnel and ICSI in large animals is not as well
established as in the mouse. Nonetheless, it allows the
introduction of very large DNA fragments favouring
proper gene expression. The efficiency of ICSI-MGT
has been improved by the use of transposons using fresh
sperm in the mouse (Moisyadi et al., 2009).
Additionally, the use of transposons together with the
injection of round spermatids has resulted in the
production of genetically modified mice (Suganuma et
al., 2005; Moisyadi et al., 2009).
Germ cell transplantation &male germline stem cells
(GSCs)
Male GSCs are unipotent adult stem cells in the
testis that self-renew and undergo differentiation to
eventually form sperm (Dym, 1994). The ability of
GSCs to transmit genetic information to the next
generation makes them an attractive vehicle for passing
on genetic modifications. The germ cell transplantation
technique developed by Brinster and coworkers in 1994
laid the foundation for using GSCs as a transgene
carrier as it demonstrated that GSCs from a donor
mouse testis, when transplanted into the seminiferous
tubules of an infertile recipient testis, were able to
colonize the recipient testis and re-established long-term
donor-derived spermatogenesis (Brinster and Avarbock,
1994; Brinster and Zimmermann, 1994).
As GSCs are difficult to maintain in vitro and
proliferate very slowly, the prevailing method of
introducing exogenous genes into GSCs relies on viral
vector-mediated transduction. The resulted genetic
modification is random insertion of the transgene into
the genome. Transduction of rodent GSCs by lentiviral
or retroviral vectors proved to be successful and resulted
in transgenic mouse and rat offspring after germ cell
transplantation (Nagano et al., 2001, 2002; Hamra et al.,
2002; Ryu et al., 2007; Takehashi et al., 2007). Viral
vectors have also been used to transduce GSCs from
large animals to make transgenic gametes and embryos
(Honaramooz et al., 2008; Zeng et al., 2013).
Due to the saftey concerns and the limited
packaging size of the viral vectors, an alternative
method, namely nucleofection has been applied to
deliver transgenes into GSCs. Nucleofection presents a
technical advancement over traditional electroporation
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Tang et al. Germline modification of domestic animals.
technology with improved transfection efficiency and
survival rate. It also allows delivery of large transgene
constructs and transfection of non-dividing cells
(Trompeter et al., 2003; Lorenz et al., 2004; Zeitelhofer
et al., 2007). Nucleofection has been used successfully
to transfect rodent and goat GSCs prior to
transplantation (Kanatsu-Shinohara et al., 2006; Izsvak
et al., 2010; Zeng et al., 2012).
There are several advantages of GSC mediated
genetic engineering. First, introduction of genetic
modifications in GSCs can circumvent problems
associated with embryo manipulation and nuclear
reprogramming (Niemann et al., 2005; Bacci, 2007;
Galli et al., 2012). Secondly, spermatogenesis in vivo
also provides a natural selection scheme to eliminate
undesired mutations that disrupt fundamental cellular
processes, and the mutations that are detrimental to
spermatogenesis. In addition, once transgenic GSCs recolonize the seminiferous tubules of the recipient testis,
they can continuously produce transgenic sperm over
the life course of the recipient. Finally, by transplanting
transduced GSCs into prepubertal recipients, the time to
production of transgenic sperm is shortened compared
to production of a transgenic founder animal by somatic
cell nuclear transfer or by ES cell-based germline
transmission (Fig. 1).
Figure 1. Approaches to generate transgenic large animals. If authentic ES cells from large animals become
available, genetic modifications can be engineered in ES cells which mediate the germline transmission of the
genetic modifications (left). The prevailing approach to make transgenic large animals is via SCNT (middle). Germ
cell transplantation using genetically modified germline stem cells (right) emerges as an alternative approach for
transmitting genetic modifications because the time required to obtain genetically modified gametes is shortened via
germ cell transplantation compared to the other two approaches.
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Tang et al. Germline modification of domestic animals.
Random integration vs. Targeted mutagenesis
The applications of genetically modified large
animals generally fall into two categories (Laible and
Alonso-Gonzalez, 2009; Aigner et al., 2010; Tan et al.,
2012): 1) over expression of an exogenous gene to
confer certain characteristics (such as a reporter), to
enhance certain traits (such as growth hormones for
animal production), or to be used as bioreactors for the
production of bioactive materials (such as Factor VIII
and alpha-1 antitrypsin). In those scenarios, the
exogenous DNA is usually integrated into the genome
in a random fashion and extensive screening of the
transgenic animals is desired for stable, prolonged and
trans-generational expression of the transgene; 2)
engineering genetic changes in a specific location in the
genome for the purpose of disease modeling, trait
modification or precise spatial/temporal expression of
an exogenous gene, or gene silencing. In this case, the
exogenous DNA carrying the desired genetic
modifications is “targeted” to a specific locus by
homologous
recombination
(Capecchi,
2005).
Alternatively, double strand (ds) breaksare created in a
specific locus by engineered nucleases and genetic
modifications are brought about as a result of DNA
repaireither by Non-Homologous End Joining (NHEJ)
or homologous recombination (HR).
Transgenesis through random integration
The majority of transgenic large animals were
produced by random integration of the exogenous
transgene into the genome. There are a few concerns
associated with random integration of the transgene.
The transgene is usually carried on a plasmid or a virus
based vector, which results in the presence of undesired
vector sequences (bacterial or viral) upon integration
into the genome. The linear transgene tends to form
concatemers upon insertion into one or more loci,
resulting in potential structural instability and transgene
silencing by the host. Random integration can also lead
to insertional mutagenesis when the transgene is
inserted into a functional gene. This represents a
particular concern on viral vectors as analysis on the
integration site of several viruses in the human genome
indicates that some viruses have integration preferences
towards active genes and their regulatory motifs
(Schroder et al., 2002; Wu et al., 2003; Berry et al.,
2006). By design, the expression of a transgene is
usually regulated by a promoter engineered in the
expression cassette; however, the site of integration may
affect the spatial and/or temporal expression of the
transgene as a result of position effects or epigenetic
silencing over time.
Compared to un-facilitated integration of
exogenous DNA into the host genome, DNA
transposons allow more efficient integration of the
exogenous transgene without the contamination of
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vector sequences and are less prone to integrate in the
form of concatemers. Those features promoted
transposon-based insertional mutagenesis to become an
important tool of large-scale functional genomics in
several species (Kumar and Snyder, 2001; Carlson et al.,
2003; Bessereau, 2006; Largaespada, 2009; Furushima
et al., 2012; Yergeau et al., 2012). The most common
transposon system used for genetic engineering is class
II DNA transposons which use a “cut-and-paste”
transposition mechanism to excise a precise DNA
segment from a DNA sequence (i.e. from a vector) and
insert it into another DNA sequence (i.e. the host
genome; Ammar et al., 2012). The transgene expression
cassette to be transposed is carried on a transposon
vector and flanked by inverted terminal repeats (ITRs)
of the transposons. The transposase enzyme, which is
provided together with the vector, recognizes the ITRs
and drives the transposition reaction.
Several transposon systems such as Sleeping
Beauty, Tol2 and piggyBac have been used as gene or
enhancer trap vectors for functional genomics as well as
for transgenesis in fly, fish and rodents (Carlson et al.,
2003; Davidson et al., 2003; Balciunas et al., 2004;
Bonin and Mann, 2004; Parinov et al., 2004;
Largaespada, 2009; Nakanishi et al., 2010). Pronuclear
injection of a transposon vector and the hyperactive
Sleeping Beauty transposase into fertilized rabbit
oocytes resulted in stable transgenic rabbits (Ivics et al.,
2014). Through the approaches such as cytoplasmic
injection into zygotes or transfection of porcine
fibroblasts followed by SCNT, the Sleeping Beauty
transposon system has also been used to generate
transgenic pigs that carry a fluorescent reporter (GFP,
YFP or Venus) ora human ApoBEC3G gene (Carlson et
al., 2011, Garrels et al., 2011, Jakobsen et al., 2011).
Genome-wide mutagenesis has been achieved in rat
germline stem cells (GSCs) by using Sleeping Beauty
(Izsvak et al., 2010). Transplantation of a polyclonal
library of transfected GSCs or individually picked
monoclonal GSC lines into the recipient rat testis
resulted in germline transmission of the mutations and
generation of knockout rat offspring.
Targeted mutagenesis through engineered nucleases
Precise gene alteration via homologous
recombination (gene targeting) in mouse ES cells and
subsequent germline transmission of the mutations
revolutionized the field of functional genetics and
became the gold standard for creating mouse models for
biomedical and pharmaceutical studies. For large
animals where germline-competent ES cells are not
readily available, production of large animals with
precise genetic modifications relies on gene targeting in
somatic cells followed by somatic cell nuclear transfer
(Laible and Alonso-Gonzalez, 2009). Due to the low
efficiency of gene targeting in somatic cells,
developmental problems associated with SCNT, and the
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Tang et al. Germline modification of domestic animals.
high cost in large animal husbandry, knockout domestic
animals available to the biomedical community are
scarce.
A recent advent of engineered chimeric
nucleases greatly advances the process of site-specific
genetic engineering in large animals. Zinc finger
nucleases (ZFN) were the first nucelases described for
this purpose (Kim et al., 1996). ZFNs are composed of
zinc finger DNA recognition domains tethered to a FokI
endonuclease domain. When two ZFN monomers
recognize and bind to their DNA target in the correct
orientation, the FokI endonuclease domains dimerize
and induce double strand (ds) breaks in DNA. The ZFNinduced ds breaks can be repaired either via nonhomologous end joining (NHEJ) or via homologous
recombination (HR; Porteus and Carroll, 2005). NHEJ is
error-prone and imperfect repairs result in insertions and
deletions (indels) at the break sites. Some indels can lead
to gene inactivation by frame shift without integration of
any exogenous sequences; however, the variable nature
of indels does not allow specific engineering of the
target sequence. Alternatively, if a homologous
sequence with desired genetic changes is provided in
trans, HR allows the changes to be incorporated into the
genome as a result of ds breaks repair.
ZFNs have been successfully used for gene
targeting in primary cells, transformed cells, stem cells
as well as iPS cells (Hockemeyer et al., 2009;
Hockemeyer and Jaenisch, 2010). Moreover, knockout
rodents and fish have been generated by embryonic
injection of ZFNs (Meng et al., 2008; Geurts et al.,
2009; Carbery et al., 2010; Cui et al., 2011). The first
application of ZFNs in domestic animals was to produce
eGFP knockout pigs by ZFN-facilitated targeting of the
eGFP gene in GFP transgenic porcine fibroblasts and
SCNT (Whyte et al., 2011). This study provided a
proof-of-principle for the application of ZFNs in genetic
engineering of domestic animals. Soon after, the same
approach was used to generate knock out pigs and cattle
(Hauschild et al., 2011; Yang et al., 2011b; Yu et al.,
2011; Bao et al., 2014; Luo et al., 2014). In some of
those studies, bi-allelic mutations were identified in
somatic cells prior to SCNT and used to produce
homozygous KO animals (Hauschild et al., 2011; Bao et
al., 2014; Luo et al., 2014). Bi-allelic modification
greatly reduces the time and cost needed to generate
homozygous KO animals given the long generation
intervals of domestic animals.
Transcription Activator-like Effector Nucleases
(TALENs) came to the center of the stage as a superior
alternative to ZFNs with their much simpler design and
assembly and boarder targeting range (Joung and
Sander, 2013). Similar to ZFNs, TALENs are sequencespecific nucleases consisting of the FokI endonuclease
domain and themodular DNA binding domain of
TALEs (Christian et al., 2010). At the target site,
TALENs create ds DNA breaks to generate genetic
Anim. Reprod., v.12, n.1, p.93-104, Jan./Mar. 2015
modifications via NHEJ or HR. TALENs have been
used for gene modification in a wide range of cell types
and species including domestic animals (Carlson et al.,
2013; Joung and Sander, 2013). Mono-allelic and biallelic modifications can be generated by TALENs
when delivered into porcine or bovine zygotes or
primary fibroblasts (Carlson et al., 2012). In addition,
large chromosome deletions and inversions can be
induced in livestock fibroblasts when two TALENs
targeting the same chromosome were delivered into
cells simultaneously (Carlson et al., 2012). Cloning with
TALENs-targeted porcine primary fibroblast resulted in
KO pigs with mono-allelic and bi-allelic mutations in
the LDL receptor gene (Carlson et al., 2012). Very
recently, microinjection of TALENs and ZFNs into pig
zygotes resulted in production of live piglets with
targeted mutations (Lillico et al., 2013). Although the
efficiency of nucleases-mediated gene targeting is
relatively high, gene mutations created by NHEJ are
unpredictable due to the nature of the repair. HRdirected repair is desired to engineer specific gene
modifications. This has recently been demonstrated in
generation of specific KO mice and pigs by
simultaneous delivery of TALENs and DNA
oligonucleotides (Tan et al., 2013; Wefers et al., 2013).
The HR-directed repair has also been achieved in pig,
goat and cattle fibroblasts with both mono-allelic and
bi-allelic modifications (Tan et al., 2013).
A recent addition to the designer nucleases
family is a RNA-guided nuclease system named
Clustered Regularly Interspaced Short Palindromic
Repeats (CRISPRs)/CRISPR-associated (Cas) 9 (Jinek
et al., 2012). The CRISPRs/Cas9 system was adopted
from the adaptive immune system of bacteria and
archaea against foreign DNA in which CRISPR RNAs
(crRNAs), with the help of trans-activating crRNA
(tracrRNA), guide the endonucleases Cas9 to induce
ds breaks in target DNA (Jinek et al., 2012). Instead of
having a DNA binding domain for DNA recognition as
in ZFNs and TALENs, CRISPRs/Ca9 uses RNAs as a
guide for site specific DNA recognition and recruits
Cas9 endonucleases for DNA cleavage. Ever since
2012, the CRISPRs/Cas9 system has been successfully
adapted for engineering mono-allelic and bi-allelic
gene modifications in cells and multicellular
organisms (Wilkinson and Wiedenheft, 2014). In
addition, multiple guide RNAs can be encoded into a
single CRISPR array to allow simultaneous multiple
gene targeting in one step (Cong et al., 2013; Ma et al.,
2014). This presents an exciting opportunity to knock
out redundant genes, multiple members of a gene
family or multiple players in signaling pathways. The
most recent breakthrough in large animal genome
modification was achieved in cynomolgus monkeys
using CRISPRs/Cas9 (Niu et al., 2014). Two genes
(Ppar-γ and Rag1) were disrupted simultaneously in
KO monkeys (Niu et al., 2014).
99
Tang et al. Germline modification of domestic animals.
Conclusion and perspectives
Genetically modified large animals provide
important models for the study of human and animal
disease, as pre-clinical platforms for the development
and testing of new treatments and hold potential to
improve efficiency and animal well-being in production
agriculture. Transgenesis through SCNT is currently the
prevailing technology to generate transgenic large
animals; however, other strategies such as genetic
modification through the male germ line provide
alternative approaches. While the majority of existing
models have involved random insertion and gain-offunction, the availability of designer nuclease
technology, when combined with SCNT or germ line
modification, now opens new avenues for targeted
modifications and generation of loss-of-function
(knockout) models.
Acknowledgments
Work from the authors’ laboratory was
supported by NSERC Discovery Grant 385837-2010
and NIH/ORIP 9 R01 OD016575-12.
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